Lidstrom: Molecular Devices Plate Reader

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Revision as of 09:13, 18 December 2013 by Janet B. Matsen (talk | contribs) (Plates)
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Our System

  • Machine = molecular devices: spectramax 190
  • Software = Softmax Pro, version 5.3. (manual)

WARNING: THIS SOFTWARE CRASHES PRETTY EASILY . Worse, any file that is open while the program crashes will be corrupted.

  • Things that cause it to crash:
    • Pressing save while data is being recorded
    • Making a new plate while data is being recorded, if the plate selected when you click to make a new plat is the plate being recorded.
    • Having files that are "too big."
      • Amanda had it freeze because the file was "too big" 12/2013. It was 1.3 MB, and froze upon saving. She gave it 5 min to save, but the program said "not responding" in the force quit box. The file produced after force quitting was broken: the data was gone when she tried to open it again. Amanda has done bigger files before, so this was a little surprising. Customer support said that it can crash if you have files that are too big. Break your runs down into different files if you are worried. Amanda also suggests giving it as long as possible; maybe it would have come back to life.
  • Save often to avoid trauma:
    • Save after each strip or chunk of strips you monitor.
    • Pressing save can be slow, but "save as" is much faster. Make many iterations of a file: file-revA.pda ... file-revK.pda
    • Consider taking screen shots often, saving often, and maybe the autosave option.
    • Note: Amanda had tried the autosave option but it was saving files in unusual locations.
  • Know that it is normal for the software to say "not responding" when saving or when making a new plate once the file gets larger.

Detection Basics

How much of my substance can I add?

  • You don't want to be above the limit of detection!

Order of reagents

Example 1:

  • add 20-40uL cell extract per well. Add ~180 uL of mix of substrate & cofactors dissolved in buffer. Adding the large volume to the small volume should yield decent mixing.

Should I run a whole plate at once or rows/columns at a time?

  • There are a few reasons you want to read the wells you load as quickly as possible. As soon as you mix all your reagents and cell extract, your reaction starts. If your assay has a non-linear reaction rate at short times (often the case) then you will lose some information at early timepoints. For example, often a reaction is fastest at small times, and slows down as the reaction proceeds and products build up.
  • Unless you know your assay is highly linear for 30+ seconds, Janet recommends you run one cell extract with every substrate/no-substrate mix per scan of the machine, then move to the next one. OR, run all of your cell extracts with one substrate/no-substrate mix at a time.

Do I need to use the crystal plate or a plastic plate?

  • NADH assays are run at 340 nm. This is right on the lower boundary for what wavelengths are suitable for our plastic so it shouldn't matter much. See discussion and data below.

Exporting Data

  • If kinetic data needs to be exported and you can't get the data exported as one column per well:
    • Go to Settings --> Preferences
    • Change the settings to Time
    • Your files should look like this:
      best export format for Softmax Pro-kinetic data
      • Note: This assay ran for < 1 minute and Microsoft excel misinterpreted the times as minutes, not seconds. This may not be a problem if you run your assay longer, but it was a problem here.
    • (from manual)

Kinetic Data

  • The machine can report Vmax in units of milli-Units/min or Units/second. What are "Units" you ask? It is just the slope. Multiply by 1,000 to get d(Absorbance)/min.


What type to use when detecting NAD/NADH

  • You can chose between the crystal plate and disposable plastic plates. The decision is a trade-off between getting (possibly) better optics and less interference at 340nm with a crystal plate, however the crystal plate builds up residue that may lead to inaccurate assay results.
  • Disposable plate materials available: (source)
    • Methacrylate cuvettes are designed for accuracy throughout the VIS-UV spectral range from 285 nm to 750 nm.
    • Poly methacrylate cuvettes are ideal for concentration measurements in the 285 nm to 750 nm spectral range.
      • The Lidstrom lab doesn't stock these, as of 11/2013.
    • Polystyrene cuvettes are designed for assays throughout the 340 nm to 750 nm visible spectral range.
      • These are the default kind of cuvette & 96 well plate stocked by the lidstrom lab as of 11/2013. Janet got excellent results using them and they are only ~$1.50/plate.
    • UV cuvettes are ideally suited for measurements at 260 nm, 280 nm and in the visible range.
  • BioTechniques shows % transmittance of polystyrine (green line) at different wavelengths:
    • % transmittance of plates used in a plate reader. Polystyrine (the default plastic) is shown in green.

Data showing NADH standards in the quartz plate and a polystyrene plate

  • This experiment shows NADH standards run with 150uL well volume. All data can be found here.
  • Time course:
    NADH standards run for 10 min in two plate materials. (data)
  • Average readings between 2.5 and 10 min:
    NADH standards ranging from 0 to ~5mM NADH. (data)
  • Zoom in to ~1mM. Note the polystyrene plate with only buffer has absorbance. This isn't important if only the slope of A340 is required, but it should be kept in mind.
    NADH standards ranging from 0 to ~5mM NADH -- zoom into ~1mM. (data)
  • Why are the slopes different?
    • It is actually surprising that the polystyrene plate has a similar slope to the quartz plate, given that the polystyrene plate is supposed to have some amount of transmittance loss.
    • Why is the slope higher for the pink line, which was calculated based on the A340 you would expect from an ideal solution? Possible explanations include inaccuracy in the mass weighed out and partial oxidation of the NADH freezer stock.

How to clean the crystal plate

From Amanda Smith 11/2013:

Use tips

  • sharpie color the bottom of wells you have used on disposable plates. This does two things: (1) prevent you from re-using a well that has been used and (2) reduce pipetting errors (adding a reagent or mix to the wrong well)